Towards application of CRISPR-Cas12a in the design of modern viral DNA detection tools (Review)

Early detection of viral pathogens by DNA-sensors in clinical samples, contaminated foods, soil or water can dramatically improve clinical outcomes and reduce the socioeconomic impact of diseases such as COVID-19. Clustered regularly interspaced short palindromic repeat (CRISPR) and its associated protein Cas12a (previously known as CRISPR-Cpf1) technology is an innovative new-generation genomic engineering tool, also known as ‘genetic scissors’, that has demonstrated the accuracy and has recently been effectively applied as appropriate (E-CRISPR) DNA-sensor to detect the nucleic acid of interest. The CRISPR-Cas12a from Prevotella and Francisella 1 are guided by a short CRISPR RNA (gRNA). The unique simultaneous cis- and trans- DNA cleavage after target sequence recognition at the PAM site, sticky-end (5–7 bp) employment, and ssDNA/dsDNA hybrid cleavage strategies to manipulate the attractive nature of CRISPR–Cas12a are reviewed. DNA-sensors based on the CRISPR-Cas12a technology for rapid, robust, sensitive, inexpensive, and selective detection of virus DNA without additional sample purification, amplification, fluorescent-agent- and/or quencher-labeling are relevant and becoming increasingly important in industrial and medical applications. In addition, CRISPR-Cas12a system shows great potential in the field of E-CRISPR-based bioassay research technologies. Therefore, we are highlighting insights in this research direction. Graphical Abstract

Introduction
It is estimated that there are ten nonillions (10 31 ) individual viruses on our planet, and more than 7000 viral genotypes have been extensively studied and described [1][2][3][4]. A virus is an infectious pathogen agent of a noncellular structure that cannot reproduce outside the host cell. Replication of the viral genome occurs only inside the living (host) cells. They are infectious agents that can infect any cellular organism (prokaryotic, eukaryotic, and archaea) [5][6][7][8][9]. Furthermore, viruses are found in almost every ecosystem on the planet. However, viruses are smaller than bacteria, making them impossible to see under a light microscope, using only electron microscopes (cryo-electron microscopes, transmission electron microscopes) or X-rays to visualize them [10]. Moreover, the origin of the viruses has not been elucidated because they do not form fossils [11], but molecular technology has been most helpful in exploring their origin and creating a classification. However, since 1892 [12], the classification of viruses has changed several times. André Lwoff, Robert Horne, and Paul Tournier [13] were the first who develop a virus classification tool based on the Linnaean hierarchical system. In 1966 when the International Committee on Viral Taxonomy (ICTV) was established, the Baltimore [14] classification system began to be used as a traditional hierarchy of viruses. In most cases, viruses can be grouped according to their genetic material: DNA or RNA.
DNA viruses (herpes viruses, smallpox viruses, adenoviruses, human papillomaviruses, pararetro viruses, Etc.) are responsible for the most significant viral infections. The genome of DNA viruses is based on deoxyribonucleic acid (DNA), and their replication, transcription, and immunization involve DNA-modifying enzymes (DNA polymerases, Reverse Transcriptase, CRISPR-Cas, Etc.) [15], initially can be present in the virus hosting cell and/or possessed by a virus [16][17][18]. Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) together with a CRISPR associated protein (Cas) and guided by short CRISPR RNA (gRNA) acts as an 'immune' and/or antiviral system of prokaryotic organisms such as bacteria and archaea [19]. However, DNA viruses are very often infecting both prokaryotic and eukaryotic microorganisms and, therefore, the genome of DNA viruses is diverse [18].
The genomes of DNA viruses, which can be singlestranded (ssDNA) or double-stranded (dsDNA), encode only a few genes (proteins) [20][21][22][23]. An infectious particle of a virus called a virion consists of a nucleic acid surrounded by a proactive layer of a capsid protein. Capsid (diameter of 20 to 300 nm) [24] is the protein envelope of the virus that encloses its genetic material. It consists of several oligomeric (repeating) structural units made from proteins called protomers. Protomers are made up of identical protein subunits called capsomeres. Viral genomes are circular, as in polar viruses, linear, and adenoviruses [25][26][27]. Most viruses control cellular mechanisms for macromolecular synthesis in the late phase of infection, directing it to synthesize large amounts of viral mRNAs and proteins rather than thousands of normal cellular macromolecules. Viruses often express proteins that modify host cell processes to maximize viral replication [28][29][30][31]. In many cases, the replication of the viral genome of most DNA viruses takes place in the cell nucleus, and here, viruses are completely dependent on host cell DNA synthesis processes. In other cases, DNA viruses from larger genomes can encode most of this cell mechanism by themselves [32]. In eukaryotes, the viral genome must pass through the cell nucleus membrane to reach these metabolic processes, and in bacteria, it must Fig. 1 Performance of basic CRISPR-Cas system: a Overview of schematic CRISPR definition; b Cas proteins are nucleases, endoribonucleases, helicases, or/and integrases: (1) single-strand DNA (break) by nuclease activity; (2) double-strand DNA (break) by nuclease activity; (3) double-strand DNA unwinding by helical activity; c Structure of Cas protein, guide RNA (crRNA) and target DNA complex; d Performance of four-part CRISPR-Cas action mechanism in the prokaryotic cell: 1. Stage-adaptation; 2-3. Stages-expression; 4. Stage-interference only enter the cell [33][34][35][36]. However, viruses use vital metabolic pathways in host cells for replication, making them challenging to eliminate from living organisms without drugs that typically cause toxicity to host cells. The most effective medical approaches to viral diseases are immune-to-infectious vaccines and antiviral drugs that selectively interfere with viral replication. Therefore, early detection of viral pathogens by sensitive bioassay methods in clinical samples, contaminated foods, soil, or water can significantly improve clinical outcomes and reduce the socioeconomic impact of viral diseases.
Quantitative real-time PCR (qPCR) [37], next-generation sequencing (NGS) [38,39], enzyme-linked immunosorbent assays (ELISA) [40] are currently the most widely used "gold standard" methods, which are applied for the detection and identification of viral DNA in clinical practice [41][42][43]. Therefore, during the recent epidemic of COVID-19 [44,45], it is especially relevant to develop a new method or to adapt/optimize other specific, sensitive, rapid, inexpensive, accurate, already applicable techniques for early detection of a nucleic acid of interest in specific and environmentally friendly methods.
Many of the applicable properties listed above are suitable for biosensors based on enzymatic reactions and electrochemical signal determination methods. Electrochemical response-based biosensing platforms are widely used due to their fast performance, affordable system, relatively simple sensing procedures, and direct determination of analytes [46]. One of the critical challenges for such a system is accuracy. However, the applicability of various enzymes combined with inorganic (silica [47], gold [48][49][50], carbon [51,52]) nanoparticles for biosensor design has been successfully evaluated in many types of research [51,[53][54][55][56][57]. These studies have shown that biosensors combined with enzymes and gold-nanomaterials increase the bioassay system's accuracy, specificity, sensitivity, and selectivity [58,59]. Hence, gold-based nanomaterials have demonstrated good performance in many applications and may be an attractive candidate for developing a CRISPR-Cas12a based system for several reasons. Gold nanoparticles are stable material, and it is easier to control particle size and composition by synthesis [60,61]. The most advantageous property of the goldbased nanomaterial is their biocompatibility with various biomolecules [62]. Therefore, gold-based nanomaterials can be used the further developments of biosensors based on DNA-and RNA-modifying enzymes.
Some reviews on CRISPR-Cas diversity, classification, and evolution have been published over the last three years [63][64][65]. In 2017, to systematize the classification, all complex encoded effector proteins and representatives have been divided into two classes (class 1 and 2), types I-VI, and more than 30 subtypes. Typical class 1 CRISPR-Cas system members are based on a complex of 4-7 Cas proteins (several Cas proteins and crRNAs bind together and form a functional endonuclease). Members this class 1 are widespread in bacteria (including hyperthermophiles) and archaea. Members of the class 2 CRISPR-Cas system use a single multidomain effector protein (uses a single Cas protein with crRNA) and are widespread only in bacteria. Unlike other classes, candidates of the CRISPR-Cas class 2 system (Cas9, Cas12, Cas13) are the most common and best-studied and are described and named as the best candidates for the development of genome editing tools suitable for applications in vivo and in vitro [66].
This review purposely shows and discusses previous research of CRISPR-Cpf1 (Cas12a) and even provides possible ideas for further development. The study addresses the attractiveness of the CRISPR-Cas12a system for simultaneous cis-(target) and trans-(non-target) DNA cleavage, sticky-end (5-7 bp) employment for the development of potential versatile electrochemical biosensing platforms (E-CRISPR) as DNA-sensors for the verification of single-stranded and double-stranded DNA virus-induced infections and the discovery of any other DNA-targets.

CRISPR-Cas action mechanism
The CRISPR [67] is a genomic region in the prokaryotic cell where genetic information about adaptive immunity is stored. The system was discovered in E. coli in 1987 [68]. More technically, the CRISPR system comprises regularly recurring palindromic sequence inserts in the genomic region (Fig. 1a). The palindromic repeats in the prokaryotic genomic region consist of about 21-40 bp, and regular DNA spacer repeats are about 20-58 bp in length [69]. In the prokaryotic cell CRISPR genomic region, Cas protein is responsible for preserving living cell genetic information [70][71][72]. In 2007 [73], several in vitro and in vivo studies have shown that the CRISPR-Cas complex acts as an antivirus system in a prokaryotic cell-it detects genetic information from foreign species (e.g., viruses) stores and destroys them in a particular and specific way. Astonishingly, in a prokaryotic cell, the mechanism of action of the antiviral system is efficient and straightforward. When the foreign species attacks the prokaryotic cell, these viruses' genome (RNA, DNA, or plasmids) is injected into the prokaryotic cell. Due to the CRISPR-Cas system, a short piece of foreign species genome information is taken and stored in the memory (locus) of prokaryotic cells, and later it can prevent the cell from repeated attacks by the same strain of viruses. Naturally, in the prokaryotic cell, protection from the foreign genome system is based on three stages of action: adaptation, expression, and interference ( Fig. 1d) [74,75].

The immunization and interference in prokaryotic cells
The primary mechanism of CRISPR-Cas action was defined by Brouns et al. [76], McGinn et al. [77], Yan et al. [78] and Siksnys et al. [79][80][81][82]. The Cas enzyme complex (which always requires Cas1 and Cas2) in the prokaryotic cell is involved in the natural metabolic process during the adaptation stage (Fig. 1d). Incredibly, in the Cas enzyme complex, Cas1 nuclease has integrase activity, and the Cas2 nuclease has endoribonuclease activity, which acts only together. This Cas complex recognizes short motifs (2-4 bp in length) adjacent to the protospacers [83][84][85]. Cas1 nuclease cleaves a piece of foreign species genome near the PAM and integrates it into the CRISPR locus, yielding a new spacer. In a prokaryotic cell, the CRISPR locus is the area where the peace of genetic information of infecting-foreign species is 'preserved' to protect cells from recurrent infections of the same infectious species), a CRISPR RNA is transcribed and processed into mature RNA (crRNA) [86,87]. During the expression stage (Fig. 1d), a CRISPR RNA (crRNA) molecule is formed. One of the DNA strands with CRISPR locus is transcribed into mRNA (described in some references as pre-crRNA) [77,88,89]. mRNA becomes long and exactly complementary to DNA strand, containing repeats of many CRISPR complementary sequences and genomic sequences of foreign species. crRNA is formed from the transcribed mRNA. However, the final composition of crRNA also differs for different Cas types (I-III types). In the expression stage (type I), crRNA consists of one repeat of the CRISPR genome and one genome of a foreign species. Technically, each repetition of the CRISPR sequence forms a loop, but each repetition of the genome sequence of a foreign species does not form a loop, and subsequently, the Cas6e and Cas6f nucleases digest the crRNA. The transactivating CRISPR RNA (tracrRNA) molecule plays a crucial role in the type II process. Technically, the tracrRNA sequence is digested with Cas9 nuclease and RNase III. In the type III process, the Cas6 nuclease directly disrupts each repeat of the CRISPR sequence and the foreign species genome sequence (Fig. 1d) [78,90]. In the interference stage (Fig. 1d), specially encoded crRNA (as a guide-RNA) is integrated into the Cas protein and forms the CRISPR-Cas complex [91]. In the combined system, the CRISPR-Cas complex contains genomic information from the foreign species recorded into crRNA that allows the foreign genome's identification, detection, and inactivation. This antiviral mechanism works in prokaryotic cells.
Following the present invention, bioengineered CRISPR-Cas systems have been implemented in industries that exploit bacterial cultures (dairy products, agriculture, Etc.) to establish the ability to protect a bacterial culture from virus attack [73,[92][93][94]. Later, in 2014, the CRISPR-Cas system was adjusted as a powerful tool for genomic research to silence and/or edit gene sequences with additional effectors in various organisms [85,[95][96][97]. Initial studies in bacterial cell lines [98,99] and mammalian cells [100][101][102][103][104] have shown that the biologically engineered CRISPR-Cas technology has future potential for correcting gene mutations. Such as malaria blocking genes in mosquitoes [105][106][107][108][109][110], genome editing in zebrafish [111][112][113], removing HIV genes [64,[114][115][116], hepatitis C virus [117] or Parkinson's disease [118]. However, this exciting progress may have unintended consequences and impacts due to 'off-target' effects, which recently are the main limitations of the CRISPR-Cas system because applied genetic corrections can have unpredictable results for future generations. Nevertheless, some developers of the CRISPR-Cas system (Emmanuelle Charpentier and Jennifer A. Doudna) have been awarded by Nobel Prize in Chemistry 2020 [119] and the CRISPR-Cas system has become a new generation of genomic engineering tools.
Furthermore, due to urgent need, chased by pandemics and pathogenic viruses has increased the demand for rapid, accurate, low-cost nucleic acid detection methods, and the studies have shown how the CRISPR-Cas (including Cas12a) combination in DNA Endonuclease-Targeted CRISPR Trans-Reporter (DETECTR) [120], one-Hour Low-cost Multipurpose highly Efficient System (HOLMES) [121] assays can be adapted to become an excellent biomedical diagnostics tool.

Potentials and limitation of CRISPR-Cas12a
A new potential for the Class 2 CRISPR-Cas system is the Cas12a member (CRISPR from Prevotella and Francisella 1), which consists of 1300 amino acid residues. This 151 kDa monomeric protein enhances the application of CRISPR systems to genomic engineering [122]. Recently, an opportunity came to follow the crystal structure variants of the Acidaminococcus sp. (AsCpf1), which was evaluated by McMahon et al. and Dong et al. [66,123] and Lachnospiraceae bacterium (LbCpf1) evaluated by Safari et al. [124], Jiménez et al. [125] and Swarts et al. [126]. Structural and functional differences in Cas9 and Cas12a were reported [127]. However, the primary key points are that since other class 2 (Cas9, Cas13) candidates Cas12a can be reprogrammed to recognize the target dsDNA sites. A single crRNA guides Cas12a and for cleavage, no tracrRNA is required [128,129]. Furthermore, Cas12a recognizes the T-rich PAM (5′-TTTV-3′) site by guided crRNA, and another essential distinguishing feature of Cas12a-crRNA is that it which interact with the phosphate backbone of the target DNA strand) region is in the middle of NUC and REC lobes. The cleavage mechanism: a cis-cleavage in RuvC domain; b cis-and trans-cleavages in RuvC domain; c E-CRISPR biosensor prototype based on CRISPR-Cas12a system: C1target dsDNA detection by cis-cleavage when target dsDNA is immobilized on AuNP as analyte. C2-target dsDNA detection by trans-cleavage when ssDNA is immobilized on AuNP and dsDNA is analyte. C3-target dsDNA detection by trans-cleavage when CRISPR-Cas12a is immobilized on AuNP and dsDNA is analyte. C4-dsDNA detection by trans-cleavage and additional effector when ssDNA is immobilized on AuNP and dsDNA as analyte. The CRISPR-Cas12a system can be fused with some newly designed enzymes like polymerases, other nucleases, or fluorescent proteins as additional effectors. Afterward, modified Cas protein in the CRISPR-Cas system can be used to transport those effectors to a specific DNA sequence for transcription, specific hydrolysis, visualization, or another practical purpose target. Together, through the examples detailed above, we have illustrated integrating CRISPR-Cas systems into different types in vivo biological sensing scenarios as well as emerging monitoring points compassionate and selective diagnostic programs determination of nucleic acids, proteins, and other small molecules can also be directed to suboptimal PAMs (5′-TTV-3′, 5′-TCTV-3′, 5′-TCCV-3′, and 5′-CCCV-3′) sites (Fig. 2a). Moreover, due to these PAM benefits or limitations, Gao and co-authors [130][131][132][133][134][135] have shown that science can modify robust evolutionary theories and adapt to alternative PAM sequences to increase its targeting range for the CRISPR-Cas12a system, but of course, with a lower cleavage efficiency rate. After the identification of the PAM site, CRISPR-Cas12a cleaves (42-44 bp) target (cis-) double-stranded DNA (dsDNA) of the target at 37 °C temperature and generates sticky ends (5-7 bp) near the PAM target site-this is another essential attribute of Cas12a (Fig. 2a). Hence, the CRISPR-Cas12a is different from other CRISPR-Cas systems identified as additional non-target (trans-) cleavage activity (Fig. 2b), and the protein-based part CRISPR-Cas12a is smaller than that of Cas9. Therefore, the formation of the CRISPR-Cas12a complex with crRNA is remarkably more uncomplicated. The complex formed is smaller, and crRNA forms only one loop. Furthermore, Cas12a has a protospacer (24 bp) and is more specific because Cas12a has a lower intrinsic tolerance for crRNA-target DNA mismatches and requires higher complementarity. The RuvC domain (lysine residue) is responsible for target cleavage, but differently from some other Cas representatives, Cas12a lacks the detectable second endonuclease domains (HNH) [136][137][138].
Direct comparison as mentioned above is unique in the way that the CRISPR-Cas12a system at the RuvC domain has additional nonspecific (trans-) single-stranded DNA cleavage activities dependent on Mg 2+ and Ca 2+ ions (Fig. 2b) [139,142,143]. Recent studies [144][145][146][147] have shown that the additional cleavage phenomenon is induced in rapid and complete cleavage of the ssDNA strand when the ssDNA sequence is not complementary to crRNA or other strand sequences. Furthermore, the cleavage doesn't relate to the dsDNA specific sequence. Technically, no additional PAM sequence is required for self-cleavage activation. After target dsDNA unwinding and cleavage during the ordinary CRISPR-Cas12a action, the RuvC domain becomes accessible, and the non-target ssDNA cleavage occurs spontaneously (Fig. 2b). The spontaneous cleavage indicates that activation of nonspecific ssDNA cleavage has happened in the presence of CRISPR-Cas12a target sequence for dsDNA recognition and cleavage (Fig. 2) [122,148]. Moreover, in 2020 Christopher W. Smith and co-authors [149] research proved that this CRISPR-Cas12a trans-cleavage is not limited to ssDNA substrates, and Cas12a-based diagnostics can be extended to ssDNA/dsDNA hybrid substrates. Several variables of NaCl (50-150 mM) concentration and fluorescently silent ssDNA/dsDNA (0-12 bp nicked) hybrid substrates were applied in the bioassays to monitor CRISPR-Cas12a cis-(target) and trans-(nontarget) activities. These studies have proved that CRISPR-Cas12a activity significantly reduced the increase in NaCl concentration. This CRISPR-Cas12a trans-ssDNAcleavage activity offers a new strategy to improve transcription and replication responses in vivo, label ssDNA, or develop faster, more sensitive and specific tools for the determination of specific DNA sequences.
In the studies by Gootenberg et al. [150], Kim et al. [151] and Doudna et al. [152], both Cpf1 orthologs (AsCpf1-from Acidaminococcus sp. and LbCpf1-from Lachnospiraceae bacterium) have been applied in both (i) genome editing in vivo and (ii) DNA assembly in vitro. AsCpf1 and LbCpf1 differences were reviewed in 2007 by Kim et. al. [153] and Verwaal et. al. [154]. However, geneediting studies in combination with CRISPR-Cas12a have shown the potential for 'self-processing' . The system can be assembled into a single and relatively simple plasmid suitable for transfection into selected cells to manipulate selected genes. Therefore, the CRISPR molecular tool can detect selected nucleic acid sequences, target gene editing, and a new protein detection strategy. These benefits and unique features increase the attractiveness of the CRISPR-Cas12a system to develop various biotechnological and bioassay research tools [155].

E-CRISPR application
Schematic comparison of Cas proteins in their native forms is detailed in publication by Patrick Schindele et al. [181]. The CRISPR-Cas9 system mediates its function through a single effector Cas9 and two small RNAs, crRNA and tracrRNA. After hybridization, the crR-NAR-tracrRNA complex binds to the Cas9 nuclease and binds to its recognition site before the PAM sequence. DNA binding is promoted by a 20 nucleotide reference sequence of crRNA. Cas9 nuclease causes a blunt-ended DSB 3 bp before the PAM sequence. The recognition of the crRNA-tracrRNA-target complex is promoted by the REC (recognition) section, the PI (PAM interacting) domain is responsible for recognizing the PAM. The DSB is mediated by the HNH and RuvC nuclease domains, with the HNH domain cleaving the target and the RuvC domain cleaving the non-target chain. The CRISPR-Cas13 system (Cas13a) mediates its function through a single effector Cas13 and a single crRNA. By combining Cas13 and crRNA, the complex binds to its recognition site on the target RNA mediated by the crRNA sequence. The catalytic site is outside the protein, directed to the surrounding solution, hence, the target RNA is cleaved away from the recognition site. The recognition of the crRNA-target complex is promoted by the REC (recognition) section, the cleavage of the target RNA is performed by the HEPN domain. The CRISPR-Cas12a system mediates its function through a single effector Cas12a and crRNA. By combining Cas12a and crRNA, the complex binds to its recognition site downstream of the PAM sequence. DNA binding is promoted by a 23 to 25 nucleotide reference sequence of crRNA. Cas12a nuclease induces a stepwise DSB distal to the PAM sequence. The recognition of the crRNA-target complex is mediated by the REC (recognition) section, the PI (PAM interacting) domain is responsible for identifying the PAM.
New revolutionary research by Lee et al. [182], Zhang et al. [155] and Dai et al. [183] demonstrates a universal and straightforward endonuclease activity monitoring method with the micro-fabricated gold-working electrode-based three-electrode system, where E-CRISPR modifies the gold-working electrode. In 2020, the invented E-CRISPR method is based on CRISPR-Cas12amediated interface and ssDNA reporter cleavage. The authors declared that the E-CRISPR system is suitable for detecting key categories (ssDNA, dsDNA) of biomolecules, providing the potential for implementation in the healthcare industry. Moreover, the unique transcleavage nuclease activity allows the use of any ssDNA sequence labeled by fluorescence signal reporter, this part of the E-CRISPR system can be easily replaced by another ssDNA with a signal reporter and applied for the determination of selected ssDNA, dsDNA, or ssRNA. Therefore, this aspect 'to make E-CRISPR-Cas12a system reprogrammable' is very attractive.
The evolution of the E-CRISPR system has been demonstrated by applying virus DNA (Human papillomavirus, Parvovirus, Dengue viruses) and proteins (Transforming Growth Factor b1 (TGF-b1) protein, Collagen, Aggrecan, and Bovine Serum Albumin) [92,[184][185][186]. Virus dsDNA and protein conjugated (immobilized) with DNA aptamer electrochemical (E-CRISPR) detection of methylene blue conjugated to gold nanoparticles (MBAuNP) have been successfully developed and performed by electrochemical signal detection. The above research demonstrated the specificity of the E-CRISPR-Cas12a system at sufficient limits of detection. However, the detected LODs depend on the length and structure of the non-target (trans-) ssDNA strand and applied detection method. Recently, higher efficiencies (of 96%, at 30 pM LOD) were achieved when the ssDNA (of 32 bp length) was constructed in the hair-pin structure. The system additionally contained an RNases inhibitor and was performed by EIS method with Fe(CN) 6 3−/4− as a mediator. If compared, the linear structure of ssDNA (32 bp) under the same reaction condition efficiency reaches 56%. Studies confirmed that the linear non-target ssDNA (18-40 bp in length) was immobilized on the carrier. Target DNA detection efficiency was achieved 30% due to electrochemical current outputs a detection method without Rnases inhibitors or crRNA modifiers. However, when comparing other biosensors based on the CRISPR-Cas12a system, the invented E-CRISPR system showed lower sensitivity due to several following issues. The system should first be well prepared for working with RNA, as RNA is highly unstable in ribonucleases in vivo (by tissue) and in vitro (environment) to achieve sensitivity in the electrochemical response. To achieve a higher sensitivity in recording electrochemical response, the system should first be well prepared. The RNA is highly unstable in the presence of ribonucleases in vivo (by tissue) and in vitro (environment), and the chemical modification of crRNA using phosphorothioate (PS), 2′-O-Methyl (2′-O-Me), 2′-Fluoro (2′-F), S-constrained ethyl (cEt) substitutions at the terminal 5′ or 3′ ends, or internal positions [187], or additional components such as RNase inhibitors should be involved into analytical system. As it is essential to eliminate RNases contamination and significantly increase metabolic stability and expression (in vivo), affinity, extend half-live of the system, mediate high levels of gene editing, or effectively determine the limit of detection by determining CRISPR-Cas12a methods. The Wei et al. [188] declare that the detection limit and the dynamic detection range of the E-CRISPR sensor can be further improved. The authors conducted more detailed comparative studies with the Cas12a and Cas9 systems to evaluate the effect of improving E-CRISPR sensor performance. The principle of the E-CRISPR sensor is the target induced conformational change of the surface signaling probe (containing an electrochemical tag), leading to the variation of the electron transfer rate of the electrochemical tag. To better understand the -trans cleavage efficiency, the authors investigated the effect of divalent cation (Mg 2+ ) concentration in an in vitro degradation solution, as the catalytic domain of RuvC is known to act on nuclease activity based on a bimetallic ion mechanism. Futhermore, enhancement of the detection signal was observed with increasing Cas12a reaction process up to 1 h. The specific and complementarity-dependent enzymatic activity of CRISPR is exploited beyond the detection limit of conventional electrochemical DNA sensors and, most importantly, the detection accuracy [182,188,189].
Firstly, gRNA should be designed to be complementary to the target following a correct PAM sequence. Secondly, the Cas protein with crRNA should be constructed appropriately to recognize the PAM sequence in target nucleic acid, and after target (cis-) cleavage, the nontarget (trans-) cleavage is activated. Due to the detection method of E-CRISPR based on non-target cleavage, the different duration time of a non-specific target cleavage differs, and the differences were investigated by Dai et al. [155] and Zhang et al. [187]. At the latest publication, a photoactive methylene blue dye, and biotin [182], were used by several authors, but it is hypothesized that some other redox probes (phenothiazines, ferrocenes, porphyrins, Etc.) can be applied to electrochemical [190] signal registration [189,190].
CRISPR-Cas biosensing systems are suitable for developing CRISPR-Cas12a point-of-care (POC) test devices with performance equivalent to or better than conventional diagnostics practices. Sensitive and rapid detection of nucleic acids with the naked eye is a new direction in analytical diagnostics. For on-site diagnostics, an ssDNA reporter labeled with a quenched green fluorescent molecule cleaved by Cas12a was introduced, and the resulting green fluorescence can be seen with the naked eye or in 485 nm light [193][194][195]. Point-of-care testing (POCT) is advantageous in terms of its ease of use, greater approachability on the user's friendly, more timely detection, and comparable accuracy and sensitivity, which could reduce the testing load on central hospitals [196,197].
The CRISPR-Cas system is programmable, modular, and a specific biological tool for genomic or tissue engineering, bioelectronics, and diagnostics [198]. The CRISPR system is an accessible and powerful tool for regulating biological sensing strategies based on a highly selective sensing mechanism as a functional response. Combining Cas protein with a graphene-based fieldeffect transistor (FET) has been reported femtomole (1.7 fM) sensitivity of designed analytical system towards target sequence. This was achieved within 15 min lasting action of the sensing system because an increasing amount of formed DNA significantly reduces the conductivity of modified FET-gate. In addition, charged phosphate groups involved in DNA structure affect the gate, and therefore current passing through FET is changing [199]. Some other authors reported that CRISPR-Cas as the programmable and modular tool could be integrated into a set of biosensors, as a nucleic acid-based system for a stem-loop enhanced sensitivity and selectivity through degradation activities [204], moreover, such systems can be applied for detecting various targets, including bacteria, viruses, cancer mutations, and others. High CRISPR-Cas potential in biological sensing technologies is constantly inspiring new research activities to develop a new generation of nucleic acid detection platforms. However, the drawback of CRISPR-Cas-based systems is related to the relatively low sensitivity of Cas protein.
However, most CRISPR-Cas bio-sensitization methods can directly detect nucleic acid targets, which is beneficial in combination with appropriate DNA-amplification methods. Therefore, the most currently used CRISPR-Cas biosensing systems rely on the target nucleic acid amplification, which improves sensitivity. Therefore, this additional amplification step can create some additional drawbacks and can make the system less robust.
In addition, CRISPR biological sensing techniques can only be used for known DNA detection sequences that may limit their application in some specific cases [54]. It should be noted that during the development of 'handson diagnostic systems' , the development of the suitable strategy to immobilize CRISPR-Cas systems on various interfaces is one of the most challenging key issues.

Conclusions and future perspectives
The evolution of bioassay methods based on E-CRISPR-Cas12a can be unambiguously extended without any amplification or purification-based steps. CRISPR-Cas12a is an attractive tool for detecting non-target (trans-) ssDNA cleavage in electrochemical signal-based systems. In addition, after the cis-cleavage sticky ends (5-7 bp) in dsDNA are formed, this effect can be efficiently exploited to evolve ultrasensitive biosensors for target DNA detection. CRISPR-Cas12a has successfully demonstrated the potential to be applied in susceptible systems suitable for determining exceptional resolution and time efficiency in conjunction with simple visual signal readings and quantitative determination. However, the CRISPR system has additional potential application, which is still not tapped within bio-electroanalytical methods. We also are predicting that in the very near future, DNA-modifying enzymes, including recently very famous CRISPR-Cas9 and CRISPR-Cas12a systems, which are recently finding many applications for various genome-editing related purposes, will find comprehensive application in the design of 'programmable DNA-and RNA-sensors.


Generation of transgene-free human induced pluripotent stem cells from erythroblasts in feeder-free conditions

Summary This protocol describes the generation and characterization of human induced pluripotent stem cells (hiPSCs) from erythroblasts. A key difference with classical protocols is the reprogramming of erythroblasts from a simple blood draw as opposed to fibroblasts/keratinocytes, which requires a biopsy. Moreover, working with erythroblasts ensures that no recombination of the TCR/BCR genes occurs, as opposed to T cells and whole peripheral blood mononuclear cells-based approaches. Last, this approach uses non-integrative episomes ensuring no integration of transgenes into the hiPSCs genome. For complete details on the use and execution of this protocol, please refer to Perriot et al. (2018).

1. Thaw Matrigel at 4 C. 2. Dilute Matrigel in cold DMEM/F12 as per dilution factor mentioned on the datasheet (variable for each Matrigel lot, final concentration of 100 mg/mL). 3. Add the diluted Matrigel to the dish in order to cover the entire surface. In this protocol, we are using 10 cm 2 dishes or wells which are coated with 1 mL of diluted Matrigel (10 mg/cm 2 ). 4. Incubate at room temperature for 60 min. Coated dishes can be kept at 37 C for 3 days. 5. Plates are ready to use after removal of the Matrigel solution, there is no need for washing the plates before use.

Institutional permissions
This study was performed according to the ethics protocol approved by our institutional review committee (approval CER-VD 2018-01622, Lausanne, Switzerland). All subjects enrolled or their legal representatives gave their written informed consent according to the study protocol.

MATERIALS AND EQUIPMENT
Human erythropoietin, SCF, IGF-1 and IL-3 are all dissolved in sterile PBS with 0.5% BSA. Dexamethasone is dissolved in ethanol. Powders are dissolved, aliquoted and stored at À20 C as displayed in the table below. After thawing, these reagents can be kept for 10 days at +4 C. Avoid freezing-thawing cycles. This section describes all the steps to isolate and freeze PBMCs from fresh blood.
1. Pour 10 mL of Ficoll-Paque PLUS in a 50 mL Faclon tube. 2. Pour 10 mL of the freshly drawn blood in another 50 mL Falcon tube and add 10 mL of PBS. Homogenize carefully the sample.
Note: If the isolation cannot be performed right after blood drawn, the blood can be kept at room temperature for a maximum of 8 h on a rocker.
3. Carefully layer the diluted blood sample on Ficoll.
CRITICAL: When layering the sample do not mix it with Ficoll but slowly dispense it on top of the Ficoll volume.
4. Centrifuge at 750 g for 20 min at room temperature, with the brake off.
Remove as much of the plasma (upper yellow layer) as possible using a 10 mL pipette, CRITICAL: Removing of the plasma should be done leaving the PBMCs layer (whitish ring) undisturbed at the interface between the plasma and the Ficoll.
5. Gently harvest the PBMC ring and transfer the cells in a new 50 mL Falcon: Add 30 mL of RPMI. Centrifuge at 550 g for 10 min at room temperature. 6. Discard the supernatant and resuspend the PBMCs in 10 mL of RPMI. Count the cells and centrifuge at 750 g for 10 min at room temperature. 7. Discard the supernatant and resuspend the cells in FBS/DMSO (90:10) at 10 million cells/mL. Distribute the cells in cryotubes between 5 to 10 million per cryotube and freeze at À80 C.
Note: After 3 days at À80 C, cells can be transferred to liquid nitrogen for long-term storage.

Erythroblast proliferation
Timing: 10 days This section describes all the steps to obtain a culture composed at more than 90% of CD71+ CD36+ cells ready for reprogramming. This step of the protocol is based on a previous study by van den Akker et al. (van den Akker et al., 2010).
8. On day 0, thaw 10 million of frozen human PBMCs in PBS + FBS 20%. a. Centrifuge 5 min, 300 g at room temperature. Remove the supernatant. b. Resuspend the cells by performing up-and-downs with a pipette in 1.5 mL of Erythroblast medium. c. Transfer the cells into one well of a low-binding 24-well plate. d. Incubate cells at 37 C 5% CO 2 . 9. On day 1, add 500 mL/well of Erythroblast medium, gently resuspend the cells with a 1000 mL pipette to homogenize the culture and split into two wells (1 mL/well). Add another 500 mL of Erythroblast medium per well. Incubate the cells at 37 C 5% CO 2 . 10. On day 4, sort CD71+ cells by MACS. This procedure is performed using CD71 microbeads (Miltenyi) as described in the Manufacturer's instructions: link. Briefly: a. Harvest all the cells and centrifuge 5 min at 300 g at room temperature. b. Discard the supernatant and resuspend the cells in 240 mL in MACS buffer. Add 60 mL of CD71 Miltenyi microbeads. Incubate for 15 min at 4 C. c. Add 6 mL of MACS buffer. Centrifuge 5 min at 300 g at room temperature. Discard the supernatant and resuspend the pellet in 500 mL of MACS buffer. d. Insert the MACS column into the MACS separator magnet. Wash column with 500 mL of MACS buffer. Add the 500 mL of CD71 labeled cells into the column. Wash with 33 500 mL of MACS buffer. e. Remove the MACS column from the separator magnet and place it in a 15 mL Falcon collection tube. Add 1 mL of MACS buffer into the MACS column, immediately flush out positive fraction by firmly applying the plunger onto the column. Collect the flow through (CD71+ cells fraction). Centrifuge the positive fraction 5 min at 300 g. 11. Discard the supernatant and resuspend the pellet in 1 mL of Erythroblast medium (you should obtain about 500 0 000-1 0 000 0 000 cells). Transfer into a new well of a low-binding 24-well plate. Incubate cells at 37 C 5% CO 2 .
Note: During the MACS procedure. Supernatant can be discarded by inverting the Falcon tube and cells are resuspended first by flicking the tube with a finger and then by up-anddown pipetting.
12. On day 5, resuspend erythroblasts by gentle flushing. Incubate cells at 37 C 5% CO 2 . 13. On day 6, if confluence is high ( Figures 1A and 1B), resuspend the cells and split into two wells (500 mL/well). Add 1 mL/well of Erythroblast medium. If confluence is not high ( Figures 1C and  1D), carefully remove 500 mL of medium (do not shake to plate, the cells must stay at the bottom of the well for medium removal) and add 1 mL of Erythroblast medium. Resuspend the cells to distribute them evenly in the well and incubate the cells at 37 C 5% CO 2 . 14. On day 8, repeat step 13.

OPEN ACCESS
STAR Protocols 3, 101620, September 16, 2022 CRITICAL: By day 11, more than 90% of cells should be double positive for CD71 and CD36 and confluence should be good ( Figure 2A). We routinely obtain more than 95% double positive cells but reprogramming can be carried on with a purity of 90% (troubleshooting 1). The enrichment of this population needs to be assessed by flow cytometry before proceeding to reprograming. Cells are resuspended by up and down pipetting, counted and around 150,000 cells are co-labeled with anti-CD71-PE and anti-CD36-FITC antibodies (standard flow cytometry staining procedure (Perriot et al., 2018)). Cells are then washed and analyzed by flow cytometry. While performing the purity check, erythroblasts should be kept in culture in the incubator. If percentage is lower than 90%, the culture can be prolonged in the same medium up to day 15 (troubleshooting 1).

Timing: 21 days
This section describes the different steps to obtain hiPSCs from erythroblasts: electroporation of erythroblasts, culture until appearance of hiPSC clones and picking of these clones. See Figure 2 for a general timeline. These steps are based on the original publications by Chou et al. (Chou et al., 2011) and Okita et al. (Okita et al., 2011).
Step 15 directly follows the purity check by flow cytometry if the purity of erythroblasts is >90%.
15. On day 11, perform electroporation of erythroblasts with the episomes allowing cell reprogramming. Before starting the procedure, coat 3 wells of a 6-well plate with Matrigel. a. Prepare the reagents for electroporation: i Adjust the concentration of the plasmids pCXLE-hOCT3/4-shp53-F, pCXLE-hSK and pCXLE-hUL to 1 mg/mL. ii Mix the three plasmids together by adding 2 mL of each plasmid.
iii Prepare the nucleofection mix using the solutions from the Human CD34+ Nucleofector kit (Lonza) as followed: 82 mL of nucleofection solution + 18 mL of supplement. b. Resuspend and count the erythroblasts. c. Collect 1 million erythroblasts and centrifuge 5 min at 300 g at room temperature.
Completely remove the supernatant. Freeze the remaining erythroblasts as described in step 7 for PBMCs.  g. Insert the cuvette into the Amaxa Nucleofector and start the T-016 program. h. Remove the cuvette. Foam must be visible on the surface of the mix indicating the electric shock happened. 16. Add 500 mL of erythroblast medium to the cuvette and transfer cell suspension into a 15 mL Falcon tube. Avoid repeated aspiration of the sample and bubble formation. 17. Use 500 mL of erythroblast medium to rinse the cuvette and add it to the cell suspension. Add 5 mL of erythroblast medium and distribute 2 mL of cell suspension in each Matrigel-coated well of the 6-well plate. Incubate at 37 C 5% CO 2 . 18. On day 13, carefully add 1 mL of erythroblast medium on the side of the well without removing medium from the plate not to disturb cells. Incubate at 37 C 5% CO 2 . 19. On day 14, carefully add 1 mL of ReproTeSR medium on the side of the well without removing medium from the plate not to disturb cells. Incubate at 37 C 5% CO 2 . 20. On day 16, repeat step 19. 21. On day 17, remove all the medium and slowly add 2 mL of ReproTeSR medium. Incubate at 37 C 5% CO 2 .
Note: The erythroblasts frozen at this stage can be used for restarting the procedure in case of failure or to be used as reference for assessing cellular identity (see quality control section).
Thawing of erythroblasts should be done as described in step 8.
CRITICAL: This medium change should be as gentle as possible to avoid detachment of erythroblasts from the plate. At this stage, some cells should start to adopt a morphology similar to epithelial or mesenchymal cells ( Figure 2B) (troubleshooting 2).
22. From day 18 onward, change medium everyday as in step 21.
Note: During the next seven days, reprogramming will continue to occur and newly generated hiPSCs will proliferate. Once hiPSCs are densely clustered together and form an identifiable colony (or clone) ( Figures 2C and 2D), they can be individually picked to generate an independent hiPSC line (troubleshooting 2).
23. Human iPSC clones are picked between day 25 and 32 depending on the emergence of hiPSC colonies. Before starting hiPSC picking, coat one 35 mm dish with Matrigel for each clone to be picked. Once coating is done, remove the Matrigel solution. a. Add 2 mL of StemMACS iPS-Brew XF medium supplemented with Y-27632 (10 mM). b. Scrap the chosen hiPSC clone with a 200 mL micropipette and aspirate the clumps as you detach them. c. Transfer the clumps to a 35 mm dish. d. Name the newly generated hiPSC line with unique ID. Cells are now at passage 1.
Note: Once a clone is picked and transferred to an individual 35 mm dish, it generates an independent clonal hiPSC line. Each hiPSC line should have a unique cell identifier. We suggest following the recommendations from the European Bank for induced Pluripotent Stem Cells (EBiSC) (Kurtz et al., 2018). The proposed format is as followed: XXXXXXi000-A. ''XXXXXX'' (combination of 2-6 capital letters) for the origin of the line (institute or lab). ''i'' for hiPSC lines or ''e'' for embryonic stem cell (ESC) lines. ''000'' (numerical 3-digit code) for the donor ID. ''A'' (capital letter) for the clone number. 24. Wash 3 times with PBS the well in which you just picked the hiPSC clone and add 1 mL of StemMACS iPS-Brew XF medium. 25. Repeat steps 16 and 17 individually as many times as needed to generate several lines.

OPEN ACCESS
CRITICAL: Human iPSC clones from one well of the 6-well plate should all be picked the same day to avoid picking twice the same clone.
CRITICAL: The picking procedure is to be done with extreme caution as you have to work over the wells without any antibiotics. Be very careful to avoid as much as possible passing over the wells in order to prevent from contamination.
Note: Several hiPSC lines should be generated per donor to ensure that any phenotype observed in downstream experiments is the reflection of an intrinsic feature coming from the donor and not a byproduct of the reprogramming procedure that would be unique to a given hiPSC line regardless of the donor. Usually, at least two lines per donor are recommended when comparing groups of donors (e.g., patients vs healthy donors).

Timing: 21 days
This section describes how to amplify hiPSCs and bank them prior use for downstream experiments.
26. The day following hiPSC picking, remove the medium of individual clones and add 2 mL of StemMACS iPS-Brew XF medium. 27. Repeat step 26 every day from Mondays to Thursdays. On Fridays, add 3.5 mL of StemMACS iPS-Brew XF medium instead of 2 mL. The medium does not need to be changed during the weekend. 28. Once the cells have reached about 80% confluence, passage them as followed: a. Remove the culture medium. b. Wash with 1 mL of PBS without Mg2+ Ca2+ c. Add 1 mL of ReLeSR solution and incubate for about 3-5 min. Monitor the detachment process under the microscope (Figure 3). The edges of the colonies should have retracted and some holes formed within the colony ( Figure 3B). d. Carefully remove ReLeSR solution from the dish. e. Add 1 mL of Complete StemMACSä iPS-Brew medium and gently spill it on the cells.
Perform a second gentle spilling on the cells. This procedure will result in detachment of hiPSCs from the plate and generates hiPSC clumps of 100-200 mm ( Figure 3C). f. Harvest the cells in a Falcon tube and complete the volume of Complete StemMACSä iPS-Brew medium to seed at a splitting ratio of 1:8 to 1:10 (2 mL of medium per dish). g. Plate the cells in 35 mm dishes and transfer the 35 mm dishes to the incubator (37 C, 5% CO 2 ). Once in the incubator, move the dishes left to right and back and forth to ensure homogenous repartition of the clumps in the dish. h. Change the medium the day after passage and routinely as in step 27.
Note: Some hiPSCs will not detach from the plate and will thus not be harvested. In particular, differentiated cells or colonies containing differentiated cells along with hiPSCs should not detach with this procedure. It is advised not trying to detach them, in order to keep a hiPSC culture as pure as possible, without differentiated cells.
CRITICAL: The optimal seeding density should be determined empirically after several trials. A too high seeding density or poor clump repartition will result in higher differentiation. Usually, a 35 mm dish, which is confluent at 80% can be split up into 10 new dishes.
29. Cells should be amplified for at least three weeks to ensure the stabilization of the cell line and episome fading.
Pause point: Human iPSCs can be frozen at this point (troubleshooting 3). For logistic reasons, we recommend to select four lines per donor to move forward with quality controls and to freeze the remaining lines early as backup in case of quality control failures.
30. For freezing, harvest hiPSCs from one dish as described in step 28. a to e. 31. Harvest the entire clump suspension and centrifuge at 300 g for 5 min at room temperature. 32. Resuspend in 2 mL of Cryostor and divide them into four cryotubes, with 500 mL per cryotube. 33. Freeze at À80 C. After 3-7 days at À80 C, cells should be transferred to liquid nitrogen for longterm storage. 34. For cell thawing, quickly thaw the cell suspension at 37 C. The medium should not be completely thawed to increase cell survival. 35. Add 0.5 mL of StemMACSä iPS-Brew medium to the vial. 36. Harvest the cells without up and down and transfer them into 4 mL of StemMACSä iPS-Brew medium. 37. Centrifuge 5 min at 300 g at room temperature. 38. Remove the supernatant and resuspend the cell pellet in 4 mL of StemMACSä iPS-Brew medium supplemented with Y-27632 (10 mM). 39. Distribute 2 mL per 35 mm Matrigel-coated dish and incubate at 37 C. The next day, change the medium and continue cell culture as described above.

Human iPSC quality controls
Timing: 3 weeks Prior to downstream use, hiPSCs need to pass several quality controls (QCs) (Figure 4). These controls are part of the good practices for hiPSC research as described by the EBiSC (Steeg et al., 2021;Sullivan et al., 2018). These QCs are absolutely necessary to ensure of the good quality of the hiPSCs. They have been regrouped in Table 1.
To generate embryoid bodies in order to assess the pluripotency of hiPSCs, follow the procedure below.
40. Remove hiPSC medium from one 35 mm dish at 80% confluence.  Figure 3B). 43. Remove accutase and add 2 mL of DMEM/F-12 medium supplemented with 20% fetal bovine serum (FBS). 44. Carefully scrap the hiPSC colonies using a cell lifter taking care of keeping the cells as clumps. 45. Transfer the hiPSC clumps into 1 well of a low-binding 6-well plate. 46. The next day, EB should have formed. Split them into 2 wells of a low-binding 6-well plate and add 1 mL of DMEM/F-12 + 20% FBS to each well. 47. Perform 2/3 of medium change every 2-3 days for 8 days. 48. On day 10 after EB formation, plate the EBs on Matrigel. At this stage, EB should have a round or oblong morphology with size between 300-600 mm. 49. Continue the culture by changing the medium every 2-3 days. 50. After 14 days, fix the cells with PFA 4% for 10 min at room temperature. Wash 3 times with PBS and stain the EBs for assessing the expression of the markers from the three embryonic germ layers (ectoderm (Pax6), mesoderm (SMA) and endoderm (AFP)).

EXPECTED OUTCOMES
The protocol described here allows obtaining between 10 to 20 hiPSC clones per donor. In addition to the protocol to generate the hiPSCs, we provide the methods to operate the necessary QCs ensuring that the hiPSCs obtained classify for downstream experiments according to the EBiSC standard. Not all clones generated need to be assessed for QCs but all clones used in any study should passed all mandatory QCs. Overall, this manuscript contains all the information to obtain fully characterized hiPSC clones free of transgenes and without BCR/TCR recombination, from a single blood draw.

LIMITATIONS
The protocol detailed here reproducibly generates hiPSCs responding to EBiSC standards, using homemade media and thus less expensive than using commercial kits. One limitation is that the protocol may not be directly transposable to generate hiPSC from fibroblasts since it starts from erythroblasts. It mostly is an advantage for institutes that have the capacity to recruit volunteers for their research given that a blood draw is easier to perform and less invasive than a skin biopsy. However, centers that cannot recruit their own volunteers need to rely on cell repositories. Since these cell banks mostly lack isolated PBMCs and usually bank fibroblasts, it may represent an obstacle for direct use of this protocol. Erythroblasts (double positive CD71+CD36+) should represent more than 90% of the cell culture. A lower proportion indicates issues in the erythroblasts amplification and may impact negatively the formation of hiPSCs. Most likely, poor erythroblast purity arise from issues during the amplification phase which may be due to two main reasons. First, poor storage of cytokines, growth factors and media. Second, too high or too low cell confluence will decrease erythroblast amplification.

Potential solution
The issue may be avoided by following some recommendations. First, to strictly respecting expiration date and storage conditions indicated by the supplier for each reagent. The complete erythroblast medium should not be kept more than a week and stored at 4 C at all time. We also recommend limiting at the maximum keeping the medium at room temperature. During culture, avoid at all cost to let the medium to turn yellow-orange.
If a low proportion of erythroblast is still obtained despite these recommendations, it is possible to further pursue the culture in the same conditions for four additional days and to reassess CD71 and CD36 expression at day 15. The proportion of erythroblasts should be good at this point.
If no changes of morphology are observed by day 24, it is to be considered that the reprogramming failed.

Potential solution
Failure in reprogramming is mostly associated with the quality of the erythroblasts or with an error in the electroporation (step 15). Before starting the electroporation, ensure that the erythroblasts are pure (see problem 1 above) and that the viability is more than 85%. Potential errors at the electroporation stage include miscalculation of the plasmids concentration, errors during mix preparation or technical error using the electroporation device. Make sure that all the parameters above are correct. An initial assessment of electroporation efficacy may be performed using a GFP plasmid. We recommend using the pCXLE-EGFP plasmid. Last, make sure to work quickly and to put back electroporated erythroblasts quickly in culture. Any delay causes cell death which may result in failure of reprogramming.
The survival of hiPSCs post thawing is low and does not allow recovering the hiPSC line.

Potential solution
Most likely, the freezing steps were not executed properly. In particular, one should avoid breaking hiPSC clumps as much as possible and work as fast as possible to ensure proper cell recovery.
Conversely, the thawing procedure should also be performed as fast as possible and avoiding breaking hiPSC clumps. One key element is to minimize the number of up and down done pipetting the cell suspension. Omission of Y-27632 at thawing will also result in lower cell survival.

RESOURCE AVAILABILITY
Lead contact Further information and requests for resources should be directed to the lead contact, Renaud Du Pasquier, renaud.du-pasquier@chuv.ch.

Materials availability
This study did not generate any unique reagents.

Data and code availability
This study did not generate any unique datasets or code.


Echinochrome A Inhibits Melanogenesis in B16F10 Cells by Downregulating CREB Signaling

Excessive increase in melanin pigment in the skin can be caused by a variety of environmental factors, including UV radiation, and can result in spots, freckles, and skin cancer. Therefore, it is important to develop functional whitening cosmetic reagents that regulate melanogenesis. In this study, we investigated the effects of echinochrome A (Ech A) on melanogenesis in the B16F10 murine melanoma cell line. We triggered B16F10 cells using α-MSH under Ech A treatment to observe melanin synthesis and analyze expression changes in melanogenesis-related enzymes (tyrosinase, tyrosinase-related protein 1 (TYRP1), and tyrosinase-related protein 2 (TYRP2)) at the mRNA and protein levels. Furthermore, we measured expression changes in the microphthalmia-associated transcription factor (MITF), CREB, and pCREB proteins. Melanin synthesis in the cells stimulated by α-MSH was significantly reduced by Ech A. The expression of the tyrosinase, TYRP1, and TYRP2 mRNA and proteins was significantly decreased by Ech A, as was that of the MITF, CREB, and pCREB proteins. These results show that Ech A suppresses melanin synthesis by regulating melanogenesis-related enzymes through the CREB signaling pathway and suggest the potential of Ech A as a functional agent to prevent pigmentation and promote skin whitening.

Introduction
Melanin pigmentation is caused by cosmetological and medical factors such as sunlight, hormonal changes, inflammation, age, skin injuries, or drugs [1][2][3][4]. If skin is exposed to sunlight, the main environmental factor regulating melanin pigmentation, melanin, absorbs the UV light contained in the sunlight to protect the skin below the dermis from UV damage and harmful factors such as reactive oxygen species and free radicals generated in the skin [5,6]. However, if melanin is excessively synthesized or pigmented as a result of these factors, spots, freckles, and even skin cancer can occur [1][2][3][4]. To prevent pigmentation and obtain a whitening effect, it is necessary to reduce the production of melanin by inhibiting the production process.
Melanin is synthesized in the melanosomes of melanocytes in the epidermis and determines skin and hair color [7]. The melanin synthesis process is regulated by a series of molecular and enzymatic reactions involving the microphthalmia-associated transcription factor (MITF) and three enzymes (tyrosinase, tyrosinase-related protein 1 (TYRP1), and tyrosinase-related protein 2 (TYRP2)) that are transcriptionally controlled by MITF in melanocytes [8][9][10]. Tyrosinase is an essential and rate-limiting enzyme involved in the initial step of the melanogenic pathway. It catalyzes the conversion of L-tyrosine to dopaquinone and of 3,4-dihydroxyphenylalanine (L-DOPA) to dopaquinone in the Raper-Mason pathway [11]. Sometimes, L-DOPA is produced by oxidation of L-tyrosine to activate tyrosinase. If dopaquinone is not converted to dopachrome by an additional enzymatic reaction, it enters the pheomelanin (one of the two main types of melanin) synthesis pathway. On the other hand, dopachrome converted from dopaquinone is entered into the eumelanin (the other type of melanin) synthesis pathway by TYRP1 and TYRP2 [11,12]. Because of the direct involvement of these proteins in melanin synthesis, tyrosinase, TYRP1, TYRP2, and MITF are attracting attention as candidates for the development of skin-whitening cosmetics.
When epidermal skin is exposed to exogenous stimuli, including UV, keratinocytes secrete the α-melanocyte-stimulating hormone (α-MSH). The secreted α-MSH binds to the melanocortin 1 receptor (MC1R) on the melanocyte surface to stimulate serial activation of upstream mediators in the CREB signaling pathway by inducing the activation of adenylate cyclase [13]. Eventually, phosphorylated CREB (pCREB), the active form of CREB, translocates into the nucleus and promotes the transcription of MITF, which induces the expression of tyrosinase, TYRP1, and TYRP2 to increase melanin synthesis [14,15].
Echinochrome A (Ech A) is a natural pigment commonly found in sea urchins; its chemical structure is 6-ethyl-2,3,5,7,8-pentahydroxy-1,4-naphthoquinone ( Figure 1a) [16]. Ech A has antioxidant, anti-inflammatory, antiviral, and chelating capabilities [17][18][19][20], is used as a therapeutic agent for a few diseases, and has been evaluated as a therapeutic candidate in in vitro and in vivo disease models [20,21]. Its water-soluble sodium salt form has been used to treat myocardial ischemia/reperfusion injury and degenerative diseases of the retina and cornea [22]. Previous studies have shown that Ech A alleviates not only inflammation and fibrosis in scleroderma due to bleomycin [23], but also atopic dermatitis-like skin lesions in NC/Nga mice [21], implying that Ech A could be a reagent for healing skin diseases.
Therefore, we hypothesized that Ech A would suppress melanogenesis and be a good candidate for preventing pigmentation and enhancing skin whitening. If Ech A affects melanogenesis as we hypothesize, it could be developed into a skin-whitening cosmetic based on a natural marine product. The B16F10 murine melanoma cell line is widely used to analyze melanogenesis-related mechanisms and develop whitening cosmetics [24][25][26]. In this study, we identified the concentration of Ech A that causes cytotoxicity in B16F10 cells and examined whether Ech A inhibits melanin biosynthesis. Furthermore, we investigated how Ech A affects the parts of melanin biosynthesis that are regulated by the CREB signaling pathway. Morphology of the cells exposed to different concentrations of Ech A for 24 h. Scale bar = 100 μm. (c) Viability of the cells exposed to Ech A. The cells were treated with different concentrations (0, 0.1, 0.5, 1, 10, 50, 100, 500, and 1000 μM) of Ech A for 24 h, and cell viability was measured using a Cell Counting Kit-8. Ech A treatments were conducted in three independent replicates to guarantee reliable results. To compare differences in cell viability between the control (0 μM) and Ech A-treated cells, one-way ANOVA was used. Note: * significantly different from the control cells (* p < 0.05 and *** p < 0.0001).

Effects of Ech A on Cell Viability
To investigate the effects of Ech A on the viability of B16F10 cells, we treated the cells with different concentrations of Ech A for 24 h. The morphology and viability of the cells exposed to 0.1-10 μM Ech A did not differ from those of the control cells (Figure 1b,c). Treatment with 50 μM Ech A induced altered morphology in the cells and a significant (p < 0.05) decrease in cell viability. Exposure to 100 μM Ech A reduced cell viability by more than 50%, indicating that that concentration is cytotoxic to B16F10 cells.

Ech A Inhibits L-DOPA Oxidation
To evaluate whether Ech A inhibits tyrosinase activity and L-DOPA oxidation in vitro, arbutin was used as the positive control [27,28]. In this study, the half-maximal inhibitory concentration (IC50) values of arbutin for inhibiting tyrosinase activity and L-DOPA oxidation were 126 ± 23.11 μM and 3279.74 ± 17.14 μM, respectively (Table S1). Based on those results, we selected 72 and 720 μM arbutin to test tyrosinase inhibition and 900 and 3670 μM arbutin to test L-DOPA oxidation inhibition as the positive controls.
When we measured how well Ech A exposure at different concentrations and times inhibited tyrosinase activity and L-DOPA oxidation, we found that Ech A barely inhibited tyrosinase activity at 0.1 and 0.5 μM, whereas arbutin inhibited tyrosinase activity in a concentration-dependent manner (Figure 2a). L-DOPA oxidation was slightly inhibited by low concentrations (0.1 and 0.5 μM) of Ech A (Figure 2b). We could not detect inhibition of tyrosinase activity or L-DOPA oxidation at Ech A concentrations higher than 0.5 μM because of the disturbance in the OD measurement caused by the brown color of the solution. (c) Viability of the cells exposed to Ech A. The cells were treated with different concentrations (0, 0.1, 0.5, 1, 10, 50, 100, 500, and 1000 µM) of Ech A for 24 h, and cell viability was measured using a Cell Counting Kit-8. Ech A treatments were conducted in three independent replicates to guarantee reliable results. To compare differences in cell viability between the control (0 µM) and Ech A-treated cells, one-way ANOVA was used. Note: * significantly different from the control cells (* p < 0.05 and *** p < 0.0001).

Effects of Ech A on Cell Viability
To investigate the effects of Ech A on the viability of B16F10 cells, we treated the cells with different concentrations of Ech A for 24 h. The morphology and viability of the cells exposed to 0.1-10 µM Ech A did not differ from those of the control cells (Figure 1b,c). Treatment with 50 µM Ech A induced altered morphology in the cells and a significant (p < 0.05) decrease in cell viability. Exposure to 100 µM Ech A reduced cell viability by more than 50%, indicating that that concentration is cytotoxic to B16F10 cells.

Ech A Inhibits L-DOPA Oxidation
To evaluate whether Ech A inhibits tyrosinase activity and L-DOPA oxidation in vitro, arbutin was used as the positive control [27,28]. In this study, the half-maximal inhibitory concentration (IC 50 ) values of arbutin for inhibiting tyrosinase activity and L-DOPA oxidation were 126 ± 23.11 µM and 3279.74 ± 17.14 µM, respectively (Table S1). Based on those results, we selected 72 and 720 µM arbutin to test tyrosinase inhibition and 900 and 3670 µM arbutin to test L-DOPA oxidation inhibition as the positive controls.
When we measured how well Ech A exposure at different concentrations and times inhibited tyrosinase activity and L-DOPA oxidation, we found that Ech A barely inhibited tyrosinase activity at 0.1 and 0.5 µM, whereas arbutin inhibited tyrosinase activity in a concentration-dependent manner (Figure 2a). L-DOPA oxidation was slightly inhibited by low concentrations (0.1 and 0.5 µM) of Ech A (Figure 2b). We could not detect inhibition of tyrosinase activity or L-DOPA oxidation at Ech A concentrations higher than 0.5 µM because of the disturbance in the OD measurement caused by the brown color of the solution.

Ech A Inhibits mRNA of Tyrosinase in B16F10 Melanoma Cells
To evaluate how well Ech A inhibited the expression of the tyrosinase, Tyrp1, and Tyrp2 genes, kojic acid was used as the positive control [29,30]. When we exposed B16F10 cells to serial concentrations (0, 12.5, 25, 50, 100, and 200 μM) of kojic acid, 12.5-50 μM kojic acid did not affect cell viability, but 100 and 200 μM kojic acid significantly decreased it ( Figure S1). Therefore, 50 μM kojic acid was selected as the positive control.
To investigate whether Ech A affects the expression of the tyrosinase, Tyrp1, and Tyrp2 mRNA, we used quantitative reverse transcription PCR (qRT-PCR) to measure the expression of those genes in the B16F10 cells exposed to different concentrations (0, 0.1, 0.5, 1, 5, and 10 μM) of Ech A, 100 nM α-MSH, or 50 μM kojic acid. Tyrosinase, Tyrp1, and Tyrp2 were significantly increased after both 24 and 48 h of exposure to α-MSH ( Figure 3). On the other hand, although the expression patterns of tyrosinase, Tyrp1, and Tyrp2 differed at some concentrations of Ech A between 24 and 48 h of exposure, the cells exposed to α-MSH and 10 μM Ech A together showed a significant decrease in tyrosinase, Tyrp1, and Tyrp2 expression at both 24 and 48 h compared with the cells exposed to only α-MSH. Therefore, we believe that 10 μM Ech A efficiently inhibits the increased expression of those genes caused by α-MSH.

Ech A Inhibits mRNA of Tyrosinase in B16F10 Melanoma Cells
To evaluate how well Ech A inhibited the expression of the tyrosinase, Tyrp1, and Tyrp2 genes, kojic acid was used as the positive control [29,30]. When we exposed B16F10 cells to serial concentrations (0, 12.5, 25, 50, 100, and 200 µM) of kojic acid, 12.5-50 µM kojic acid did not affect cell viability, but 100 and 200 µM kojic acid significantly decreased it ( Figure S1). Therefore, 50 µM kojic acid was selected as the positive control.
To investigate whether Ech A affects the expression of the tyrosinase, Tyrp1, and Tyrp2 mRNA, we used quantitative reverse transcription PCR (qRT-PCR) to measure the expression of those genes in the B16F10 cells exposed to different concentrations (0, 0.1, 0.5, 1, 5, and 10 µM) of Ech A, 100 nM α-MSH, or 50 µM kojic acid. Tyrosinase, Tyrp1, and Tyrp2 were significantly increased after both 24 and 48 h of exposure to α-MSH ( Figure 3). On the other hand, although the expression patterns of tyrosinase, Tyrp1, and Tyrp2 differed at some concentrations of Ech A between 24 and 48 h of exposure, the cells exposed to α-MSH and 10 µM Ech A together showed a significant decrease in tyrosinase, Tyrp1, and Tyrp2 expression at both 24 and 48 h compared with the cells exposed to only α-MSH. Therefore, we believe that 10 µM Ech A efficiently inhibits the increased expression of those genes caused by α-MSH.

Ech A Suppresses Melanin Production
To investigate the effects of Ech A on melanin synthesis, we measured the melanin concentration in the B16F10 cells that had been exposed to different concentrations of Ech A, α-MSH (100 nM), or kojic acid (50 µM). The cells exposed to α-MSH alone and α-MSH + Ech A (0.1, 0.5, and 1 µM) showed a significant increase in melanin concentration compared with the control cells (Figure 4a), implying that these concentrations of Ech A did not inhibit the melanin synthesis stimulated by α-MSH. On the other hand, the α-MSH + Ech A (5 and 10 µM) treatments induced a significant decrease in melanin synthesis in B16F10 cells compared with the cells exposed to α-MSH alone. In fact, 10 µM Ech A exhibited the same melanin production inhibitory effect as kojic acid, a direct inhibitor of tyrosinase. Furthermore, when the cells were treated with different concentrations of Ech A, the brown color of the cell pellets blurred due to the decrease in melanin production as the concentration of Ech A increased (Figure 4b). , and Tyrp2 mRNA in the B16F10 cells exposed to Ech A. Ech A or kojic acid (KA, positive control) was administered to the cells 1 h before α-MSH treatment, and then the cells were incubated for 24 and 48 h. The expression of the tyrosinase, Tyrp1, and Tyrp2 mRNA was analyzed using qRT-PCR after extracting total RNA. (a-c) Expression changes of the tyrosinase, Tyrp1, and Tyrp2 mRNA in the cells exposed to Ech A or KA for 24 h. (d-f) Expression changes of the tyrosinase, Tyrp1, and Tyrp2 mRNA in the cells exposed to Ech A or KA for 48 h. Note: * significantly different from the control cells (p < 0.05); # significantly different from the cells exposed only to α-MSH (p < 0.05).

Ech A Suppresses Melanin Production
To investigate the effects of Ech A on melanin synthesis, we measured the melanin concentration in the B16F10 cells that had been exposed to different concentrations of Ech A, α-MSH (100 nM), or kojic acid (50 μM). The cells exposed to α-MSH alone and α-MSH + Ech A (0.1, 0.5, and 1 μM) showed a significant increase in melanin concentration compared with the control cells (Figure 4a), implying that these concentrations of Ech A did not inhibit the melanin synthesis stimulated by α-MSH. On the other hand, the α-MSH + Ech A (5 and 10 μM) treatments induced a significant decrease in melanin synthesis in B16F10 cells compared with the cells exposed to α-MSH alone. In fact, 10 μM Ech A exhibited the same melanin production inhibitory effect as kojic acid, a direct inhibitor of tyrosinase. Furthermore, when the cells were treated with different concentrations of Ech A, the brown color of the cell pellets blurred due to the decrease in melanin production as the concentration of Ech A increased (Figure 4b). , and Tyrp2 mRNA in the B16F10 cells exposed to Ech A. Ech A or kojic acid (KA, positive control) was administered to the cells 1 h before α-MSH treatment, and then the cells were incubated for 24 and 48 h. The expression of the tyrosinase, Tyrp1, and Tyrp2 mRNA was analyzed using qRT-PCR after extracting total RNA. (a-c) Expression changes of the tyrosinase, Tyrp1, and Tyrp2 mRNA in the cells exposed to Ech A or KA for 24 h. (d-f) Expression changes of the tyrosinase, Tyrp1, and Tyrp2 mRNA in the cells exposed to Ech A or KA for 48 h. Note: * significantly different from the control cells (p < 0.05); # significantly different from the cells exposed only to α-MSH (p < 0.05).

Ech A Inhibits Melanin Synthesis Through the CREB Signaling Pathway
We investigated the expression of the proteins related to melanin synthesis and the CREB signaling pathway after exposing B16F10 cells to Ech A, α-MSH, or kojic acid. The TYRP1 and TYRP2 proteins were significantly increased by α-MSH treatment alone, and that increase was reversed by cotreatment with Ech A, showing that Ech A decreased protein production, especially of TYRP2, in a dose-dependent manner (Figure 5a,b). Tyrosinase

Ech A Inhibits Melanin Synthesis through the CREB Signaling Pathway
We investigated the expression of the proteins related to melanin synthesis and the CREB signaling pathway after exposing B16F10 cells to Ech A, α-MSH, or kojic acid. The TYRP1 and TYRP2 proteins were significantly increased by α-MSH treatment alone, and that increase was reversed by cotreatment with Ech A, showing that Ech A decreased protein production, especially of TYRP2, in a dose-dependent manner (Figure 5a,b). Tyrosinase was also decreased significantly by Ech A, with the highest decrease after exposure to 10 µM Ech A. In addition, pCREB expression in the cells exposed to both α-MSH and Ech A (5 or 10 µM) was significantly lower than in the cells exposed to only α-MSH and the control cells and similar to the level observed in the cells exposed to both α-MSH and 50 µM kojic acid (Figure 5a,b). Furthermore, the total CREB expression was significantly lower in the cells exposed to both α-MSH and 10 µM Ech A than it was in those exposed to only α-MSH and in the control cells. Because we could not detect the expression of MITF in whole cells, we measured its expression in the nucleus. The cells exposed to α-MSH and Ech A (0.1, 0.5, 1, or 5 µM) together showed a higher MITF expression than those exposed only to α-MSH and the control cells, whereas treatment with both 10 µM Ech A and α-MSH induced a decrease in MITF expression in the nucleus (Figure 5c,d). Therefore, we believe that Ech A inhibits melanin synthesis through the CREB signaling pathway.

Discussion
Melanin absorbs UV light from sunlight to protect the skin from UV damage, and it scavenges reactive oxygen species by absorbing harmful external factors. Thus, exposing skin to UV augments melanin production and pigmentation. Synthesis and pigmentation of excessive melanin cause spots, freckles and even skin cancer. Because people use functional cosmetics to whiten their skin, the interest in the study of melanogenesis and development of whitening cosmetics is growing. Based on previous observations that Ech A not only has antioxidative and anti-inflammatory effects, but also suppresses UVB-induced photoaging [16,31,32], we investigated the effects of Ech A on melanogenesis in

Discussion
Melanin absorbs UV light from sunlight to protect the skin from UV damage, and it scavenges reactive oxygen species by absorbing harmful external factors. Thus, exposing skin to UV augments melanin production and pigmentation. Synthesis and pigmentation of excessive melanin cause spots, freckles and even skin cancer. Because people use functional cosmetics to whiten their skin, the interest in the study of melanogenesis and development of whitening cosmetics is growing. Based on previous observations that Ech A not only has antioxidative and anti-inflammatory effects, but also suppresses UVBinduced photoaging [16,31,32], we investigated the effects of Ech A on melanogenesis in melanocytes and examined the potential of Ech A as a whitening cosmetic ingredient.
When we evaluated the cytotoxicity of Ech A in B16F10 cells, we found that Ech A did not cause cytotoxicity at up to 10 µM, but 50 and 100 µM Ech A decreased cell viability. Some previous studies reported that cytotoxicity was induced at concentrations different from those in this study [20,33]. One study reported that Ech A did not affect the viability of mouse embryonic stem cells at up to 500 µM and that up to 50 µM Ech A enhanced cardiomyocyte differentiation from embryonic bodies [33]. Another study reported that exposure to 0.5-10 µM Ech A significantly increased cell viability in human cardiac progenitor cells, and 50 µM Ech A had no effect on cell viability [20]. However, above 100 µM, Ech A significantly decreased cell viability. On the basis of previous studies and our results, we think that the concentration of Ech A that affects cell viability differs by cell type.
Melanin is mainly synthesized in two types, pheomelanin and eumelanin [11,12]. Pheomelanin varies from yellow to reddish-brown color, while eumelanin varies from light brown to black color. Tyrosinase, the rate-limiting enzyme in melanogenesis, participates in the early stages (conversion of L-tyrosine to dopaquinone and oxidation of L-DOPA to dopaquinone) of melanin synthesis [11]. In this study, in vitro tyrosinase activity and L-DOPA oxidation were evaluated to validate the potential of Ech A as an anti-melanogenic agent. Compared with the inhibition of tyrosinase activity and L-DOPA oxidation caused by arbutin, Ech A showed minimal inhibitory effects. The Ech A solution is dark brown, so we could not detect signals at concentrations higher than 0.5 µM Ech A. Therefore, it is necessary to use other experimental methods to further identify whether Ech A regulates melanin synthesis.
Arbutin, which we used as the positive control, is a cosmetic ingredient used for whitening by inhibition of tyrosinase [34,35]. However, its use is limited because of decomposition or discoloration due to its poor stability, occurrence of an off-flavor, unclear side effects, and safety issues. Therefore, a whitening agent that directly scavenges tyrosinase and inhibits the formation of melanogenesis-related enzymes at the transcription and translation levels is needed. TYRP1 and TYRP2 regulate eumelanin synthesis [11,12]. Therefore, we investigated whether Ech A affects the expression of the tyrosinase, Tyrp1, and Tyrp2 mRNA and found that the expression of those genes triggered by α-MSH was decreased after both 24 and 48 h of exposure to Ech A. In particular, treatment with 10 µM Ech A reduced the expression of the three genes to the levels expressed in the control cells. Taken together, our results show that Ech A controls melanogenesis by inhibiting the gene expression of three enzymes (tyrosinase, TYRP1, TYRP2). In this study, 5 and 10 µM Ech A inhibited α-MSH-stimulated melanin synthesis in B16F10 cells. In particular, the melanin concentration in the cells exposed to 10 µM Ech A was equal to the melanin concentration in the cells exposed to 50 µM kojic acid, the control inhibitor of melanin synthesis. Therefore, our results suggest that Ech A can inhibit melanin synthesis at a lower concentration than kojic acid.
Several signaling pathways, including CREB, are involved in melanin synthesis [28]. Keratinocytes exposed to UV secrete α-MSH, which binds MC1R on the melanocyte surface. The α-MSH/MC1R complex triggers intracellular cAMP production, which activates the CREB, JNK, MAPK/ERK, and p38 MAPK pathways [28,[36][37][38]. These signaling pathways promote MITF expression and activation, followed by increased expression of downstream proteins (tyrosinase, TYRP1, and TYRP2) and increased melanin synthesis. In this study, Ech A significantly inhibited the expression of the pCREB and CREB stimulated by α-MSH. In fact, the expression of CREB and pCREB in the cells treated with 10 µM Ech A was lower than that in the control cells. The expression of the tyrosinase, TYRP1, and TYRP2 proteins stimulated by α-MSH was also inhibited by Ech A, and 5 and 10 µM Ech A suppressed their expression with and without α-MSH stimulation. When we measured the amount of the MITF protein in the nucleus (because MITF is an important nuclear transcription factor modulating melanin synthesis [39]), we found that 10 µM Ech A effectively prevented the expression of MITF triggered by α-MSH. However, cotreatment with 0.1 µM Ech A and α-MSH tended to increase the expression of MITF compared with the cells exposed to α-MSH alone. Because 10 µM Ech A alone significantly reduced MITF expression, we think that either a low concentration of Ech A could not inhibit α-MSH stimulation or that it was an experimental error. In addition to Ech A, other natural products obtained from marine animals and plants have been investigated as potential skin-whitening agents [40,41]. Chen et al. [40] found that 196 mM chitosan from crustaceans inhibited the expression of the MITF, tyrosinase, TYRP1, and TYRP2 proteins and the tyrosinase activity triggered by α-MSH in B16F10 cells, so they suggested chitosan as a potential skin-whitening agent. Ech A is also a marine product that inhibits the expression of the MITF, tyrosinase, TYRP1, and TYRP2 proteins and the tyrosinase activity triggered by α-MSH at a concentration of 10 µM; therefore, we argue that Ech A would be more effective than chitosan for suppressing melanin synthesis. Taken together, our results suggest that Ech A is a strong inhibitor of melanogenesis.

Cell Culture
The B16F10 murine melanoma cell line was purchased from the Korean Cell Line Bank (KCLB) (Seoul, Korea). The cells were cultured in Dulbecco's modified Eagle's medium with 10% FBS and 100 units/ml penicillin plus 100 mg/ml streptomycin at 37 • C with 5% CO 2 according to the KCLB guideline. After the cells reached 70-80% confluence, they were subcultured using 0.25% trypsin/EDTA (Thermo Fisher Scientific, Waltham, MA, USA).

Cell Cytotoxicity Assay
To investigate the cytotoxicity of Ech A, cell viability was assayed using a Cell Counting Kit-8 (CCK-8; Dojindo, Kumamoto, Japan) according to the manufacturer's instructions. Prior to Ech A treatment, B16F10 cells were seeded in a 96-well plate at a density of 10,000 cells per well and incubated at 37 • C and 5% CO 2 for 24 h. Then, the cells were exposed to a fresh serum-free medium with or without different concentrations of Ech A and incubated at 37 • C and 5% CO 2 for another 24 h. At that time, CCK-8 solution was added to the plate containing the cells, and the plate was incubated at 37 • C for 2 h. Absorbance was measured at 450 nm using a microplate reader (Power Wave XS2, BioTeck, Winooski, VT, USA).

Inhibition of Tyrosinase Activation and L-DOPA Oxidation
The in vitro tyrosinase inhibition assay was performed using previously described methods [42]. The in vitro L-DOPA oxidation inhibition assay was also performed using previously described methods [43]. For the reaction mixture, 150 µL of PBS (0.1 M, pH 6.5), 20 µL of serially diluted Ech A (0, 0.1, 0.5, 1, 5, 10, and 50 µM), 20 µL of mushroom tyrosinase (2000 U/mL) (Sigma-Aldrich, St. Louis, MO, USA), and 10 µL of L-DOPA (60 µM) (Sigma-Aldrich, St. Louis, MO, USA) were added to a 96-well plate that was then incubated at 37 • C for 10-15 min, when the optical density of the wells was measured at 475 nm. The concentration of Ech A at which half the original L-DOPA oxidation was inhibited (IC 50 ) was determined using two concentrations (900 and 3670 µM) of arbutin as the positive controls.

Melanin Synthesis Assay
B16F10 cells were seeded in a 6-well plate at a density of 50,000 cells per well and incubated at 37 • C and 5% CO 2 for 24 h. Then, the culture medium was exchanged with a fresh phenol red-free growth medium containing Ech A (0, 0.1, 0.5, 1, 5, and 10 µM) or 50 µM kojic acid and the plate was incubated again at 37 • C and 5% CO 2 . After 1 h of incubation, 100 nM α-MSH (Sigma-Aldrich, St. Louis, MO, USA) was added, and the plate was incubated at 37 • C and 5% CO 2 for an additional 47 h. The cultured cells were harvested, counted, and lysed with 1 N NaOH containing 10% dimethyl sulfoxide at 80 • C for 10 min. The cell lysate was centrifuged at 13,000× g for 2 min, and the total protein concentration of the supernatant was measured. The supernatant was transferred to a 96-well plate, and the amount of melanin was measured at 490 nm. The melanin content of the total protein was calculated per test condition and is presented as a percentage of the control.

Quantitative RT-PCR
B16F10 cells were incubated with a growth medium containing Ech A (0, 0.1, 0.5, 1, 5, and 10 µM) or 50 µM kojic acid at 37 • C and 5% CO 2 for 1 h. Then, 100 nM α-MSH was added, and the cells were incubated at 37 • C and 5% CO 2 for an additional 23 h or 47 h. To analyze the expression of the Tyr, Tyrp1, Tyrp2, and Mitf mRNA, we performed qRT-PCR. The cells exposed to Ech A or kojic acid for 24 h and 48 h were harvested, and total RNA was extracted using a Trizol reagent (Thermo Fisher Scientific, Waltham, MA, USA). The extracted total RNA was reverse-transcribed with a TOPscript TM cDNA Synthesis Kit (Enzynomics, Seoul, Korea). For quantitative PCR, 5 µL of TB Green Premix Ex Taq (Takara Bio, Otsu, Japan), 5 pmol of the forward primer, 5 pmol of the reverse primer, and 1 µL of cDNA were added with water to a final volume of 25 µL. The mixture was amplified for 35 cycles using a CFX96 Touch Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA). The cycle number at which a statistically significant increase in each gene was first detected (threshold cycle, Ct) was normalized to the Ct for Gapdh. The relative expression differences between the genes were calculated using the 2 −∆∆CT method [44]. The primers used for gene amplification are presented in Table 1.

Western Blot Analysis
The cells cultured under the same conditions as in the melanin synthesis assay were lysed with a PRO-PREP protein extraction solution (iNtRON Biotechnology, Gyeonggi, Korea) containing protease and phosphatase inhibitors (Quartett, Berlin, Germany) and incubated at 4 • C for 20 min. The cell lysates were centrifuged at 13,000× g and 4 • C for 5 min, and the supernatants were transferred into fresh tubes. The extracted protein concentration was measured using a Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA, USA) according to the manufacturer's instructions. The proteins were denatured by heating, separated by SDS-PAGE, and transferred to PVDF membranes (Millipore, Billerica, MA, USA). The membranes were blocked with a TBS-T buffer containing 5% nonfat skim milk at room temperature for 1 h. The membranes were incubated with primary antibodies overnight at 4 • C: anti-pCREB (Ser133) (#9196, Cell Signaling Technology, Berkeley, CA, USA), anti-CREB (#9197, Cell Signaling Technology, Berkeley, CA, USA), anti-GAPDH (sc-25778, Santa Cruz Biotechnology, Santa Cruz, CA, USA), anti-tyrosinase (T311) (sc-20035, Santa Cruz Biotechnology, Santa Cruz, CA, USA), anti-TYRP1 (G-9) (sc-166857, Santa Cruz Biotechnology, Santa Cruz, CA, USA), and anti-TYRP2 (C-9) (sc-74439, Santa Cruz Biotechnology, Santa Cruz, CA, USA). After the membranes were washed with the TBS-T buffer, they were incubated with HRP-conjugated secondary antibodies at room temperature for 1 h. The membrane-bound proteins were visualized using a Pierce TM ECL Western Blotting Substrate (Thermo Fisher Scientific, Rockford, IL, USA). To detect the MITF and histone 3 proteins, the total nuclear proteins were prepared with NE-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific, Rockford, IL, USA) according to the manufacturer's instructions. The protein concentration was measured using a Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA, USA), and then the experimental procedure just described was performed. The primary antibodies used were anti-MITF (sc-515925, Santa Cruz Biotechnology, Santa Cruz, CA, USA) and anti-histone H3 (D1H2) (#4499, Cell Signaling Technology, Berkeley, CA, USA).

Statistical Analysis
Statistical analyses were conducted using the GraphPad Prism 9 software (San Diego, CA, USA). All the data obtained from the cell toxicity assay, qRT-PCR, melanin synthesis assay, and Western blot analysis are expressed as the means ± standard deviation (SD). The differences between the cells from different conditions obtained from the cell toxicity assay, melanin synthesis assay, qRT-PCR, and Western blot analysis were analyzed with one-way ANOVA followed by the HSD test; p < 0.05 was considered statistically significant.

Conclusions
This study demonstrated that Ech A inhibits melanogenesis and its underlying mechanism, indicating it as a potential skin-whitening agent for functional cosmetics.
Supplementary Materials: The following supporting information can be downloaded at https:// www.mdpi.com/article/10.3390/md20090555/s1, Figure S1: Cytotoxicity of B16F10 cells in response to kojic acid; Table S1: Inhibitory effects of arbutin on the activity of tyrosinase and oxidation of L-DOPA.

Conflicts of Interest:
The authors declare no conflict of interest.